Methods of identifying anti-viral agents

ABSTRACT

The present invention provides methods of identifying candidate anti-viral agents.

CROSS-REFERENCE

This application claims the benefit of U.S. Provisional Patent Application No. 60/814,229, filed Jun. 16, 2006, which application is incorporated herein by reference in its entirety.

STATEMENT REGARDING FEDERALLY SPONSORED RESEARCH

This invention was made with government support under contact F32A150296 awarded by the National Institutes of Health. The Government has certain rights in this invention.

BACKGROUND

The error-prone replication of RNA viral genomes makes them notorious for their ability to evolve resistance to selective agents rapidly and effectively. For cytoplasmic positive-strand RNA viruses such as poliovirus, other picornaviruses such as foot-and-mouth disease virus, and more distantly related flaviviruses such as Dengue and West Nile viruses, an infection started by a single genome can quickly become heterogeneous, even in the first infected cell. Therefore, a progeny genome containing a newly generated mutation that could confer a selective advantage must replicate and package in the context of an essentially polyploid infection in order to propagate.

There is a need in the art for anti-viral agents that, when administered, do not give rise to drug-resistant virus in the early stages of viral infection.

LITERATURE

-   Herskowitz (1987) Nature 329:219-222; Crowder and Kirkegaard (2005     July) Nat. Genet. 37(7):701-9 (Epub 2005 Jun. 19).

SUMMARY OF THE INVENTION

The present invention provides methods of identifying candidate anti-viral agents.

BRIEF DESCRIPTION OF THE DRAWINGS

FIGS. 1A-D depict a dominant inhibitor screen for capsid-coding genome regions.

FIGS. 2A-C depict the effect of mutations in 3D polymerase on the yield of wild-type virus during co-transfection.

FIGS. 3A-C depict a dominant inhibitor screen in the CRE and 3B-coding regions.

FIGS. 4A-E depict a dominant inhibitor screen for 2A proteinase and VP1-2A cleavage site mutant alleles.

FIGS. 5A-F depict RNA replication or translation requirements for dominance of mutant poliovirus alleles.

FIGS. 6A-C depict superinfections of wild-type and temperature-sensitive polioviruses.

FIGS. 7A-E depict co-infections of drug-sensitive and drug-resistant viruses.

FIG. 8 is a schematic depiction of an exemplary assay.

FIGS. 9-19 provide amino acid sequences of poliovirus type 1 (Mahoney) proteins VP2, VP3, VP1, 2A, 2B, 2C, 3A, 3B, 3C, 3D, and VP4, respectively.

DEFINITIONS

The terms “polynucleotide” and “nucleic acid”, used interchangeably herein, refer to a polymeric forms of nucleotides of any length, either ribonucleotides or deoxynucleotides. Thus, this term includes, but is not limited to, single-, double-, or multi-stranded DNA or RNA, genomic DNA, cDNA, DNA-RNA hybrids, or a polymer comprising purine and pyrimidine bases or other natural, chemically or biochemically modified, non-natural, or derivatized nucleotide bases.

The terms “peptide,” “oligopeptide,” “polypeptide,” “polyprotein,” and “protein” are used interchangeably herein, and refer to a polymeric form of amino acids of any length, which can include coded and non-coded amino acids, chemically or biochemically modified or derivatized amino acids, and polypeptides having modified peptide backbones.

“Recombinant,” as used herein, means that a particular DNA sequence is the product of various combinations of cloning, restriction, and/or ligation steps resulting in a construct having a structural coding sequence distinguishable from homologous sequences found in natural systems. Generally, DNA sequences encoding the structural coding sequence can be assembled from cDNA fragments and short oligonucleotide linkers, or from a series of oligonucleotides, to provide a synthetic gene which is capable of being expressed in a recombinant transcriptional unit. Such sequences can be provided in the form of an open reading frame uninterrupted by internal nontranslated sequences, or introns, which are typically present in eukaryotic genes. Genomic DNA comprising the relevant sequences could also be used. Sequences of non-translated DNA may be present 5′ or 3′ from the open reading frame, where such sequences do not interfere with manipulation or expression of the coding regions. Thus, e.g., the term “recombinant” polynucleotide or nucleic acid refers to one which is not naturally occurring, or is made by the artificial combination of two otherwise separated segments of sequence. This artificial combination is often accomplished by either chemical synthesis means, or by the artificial manipulation of isolated segments of nucleic acids, e.g., by genetic engineering techniques. Such is usually done to replace a codon with a redundant codon encoding the same or a conservative amino acid, while typically introducing or removing a sequence recognition site. Alternatively, it is performed to join together nucleic acid segments of desired functions to generate a desired combination of functions.

By “construct” is meant a recombinant nucleic acid, generally recombinant DNA, that has been generated for the purpose of the expression of a specific nucleotide sequence(s), or is to be used in the construction of other recombinant nucleotide sequences.

Similarly, a “recombinant polypeptide” or “recombinant polyprotein” refers to a polypeptide or polyprotein which is not naturally occurring, or is made by the artificial combination of two otherwise separated segments of amino acid sequences. This artificial combination may be accomplished by standard techniques of recombinant DNA technology, such as described above, i.e., a recombinant polypeptide or recombinant polyprotein may be encoded by a recombinant polynucleotide. Thus, a recombinant polypeptide or recombinant polyprotein is an amino acid sequence encoded by all or a portion of a recombinant polynucleotide.

Before the present invention is further described, it is to be understood that this invention is not limited to particular embodiments described, as such may, of course, vary. It is also to be understood that the terminology used herein is for the purpose of describing particular embodiments only, and is not intended to be limiting, since the scope of the present invention will be limited only by the appended claims.

Where a range of values is provided, it is understood that each intervening value, to the tenth of the unit of the lower limit unless the context clearly dictates otherwise, between the upper and lower limit of that range and any other stated or intervening value in that stated range, is encompassed within the invention. The upper and lower limits of these smaller ranges may independently be included in the smaller ranges, and are also encompassed within the invention, subject to any specifically excluded limit in the stated range. Where the stated range includes one or both of the limits, ranges excluding either or both of those included limits are also included in the invention.

Unless defined otherwise, all technical and scientific terms used herein have the same meaning as commonly understood by one of ordinary skill in the art to which this invention belongs. Although any methods and materials similar or equivalent to those described herein can also be used in the practice or testing of the present invention, the preferred methods and materials are now described. All publications mentioned herein are incorporated herein by reference to disclose and describe the methods and/or materials in connection with which the publications are cited.

It must be noted that as used herein and in the appended claims, the singular forms “a,” “and,” and “the” include plural referents unless the context clearly dictates otherwise. Thus, for example, reference to “an RNA virus” includes a plurality of such viruses and reference to “the anti-viral agent” includes reference to one or more anti-viral agents and equivalents thereof known to those skilled in the art, and so forth. It is further noted that the claims may be drafted to exclude any optional element. As such, this statement is intended to serve as antecedent basis for use of such exclusive terminology as “solely,” “only” and the like in connection with the recitation of claim elements, or use of a “negative” limitation.

The publications discussed herein are provided solely for their disclosure prior to the filing date of the present application. Nothing herein is to be construed as an admission that the present invention is not entitled to antedate such publication by virtue of prior invention. Further, the dates of publication provided may be different from the actual publication dates which may need to be independently confirmed.

DETAILED DESCRIPTION

The present invention provides methods of identifying candidate anti-viral agents. The invention is based at least on part on the discovery that choices of anti-viral drug target can be made so that drug-sensitive viral genomes dominantly inhibit the outgrowth of drug-resistant viral genomes in a cell. The ability of relatively unfit viruses to inhibit growth of viruses with increased fitness derives from, for example, the intracellular amplification of positive-strand RNA-viral genomes, their translation into large polyproteins, and the higher-order oligomerization of several of their protein products.

Accordingly, the invention is based in part on the concept that certain viral products, when defective, can dominantly interfere with growth of non-defective viruses, making these viral products good drug targets. Interaction of an anti-viral drug with a viral product essentially renders that viral product “defective” in function. Thus, if a viral product that, when defective due to mutation or drug interaction, is targeted by an anti-viral drug, then upon such viral products will dominantly interfere with other viruses in the cells in the presence of the drug. Stated differently, if defects similar to those introduced by a dominant mutations can be mimicked by antiviral drugs, the defective drug-sensitive viral genomes and products will dominantly interfere with the intracellular growth of any resistant mutant genomes that might arise.

Accordingly, drug sensitive viruses (i.e., viruses that produce a viral drug target that, upon drug interaction, dominantly interferes with other viruses) will be dominant over drug-resistant variants which contain a mutation in the targeted viral product. Therefore, as mutant, drug-resistant viruses arise in the cell, the presence of other viruses in the same cell that are still drug-sensitive will inhibit the growth of the drug-resistant virus. As such, this strategy will at least delay the emergence of drug-resistant viruses. This concept was borne out in experiments conducted with poliovirus, a positive-strand RNA virus.

Suitable drug targets were identified by introducing mutations into the poliovirus genome, generating variant, non-viable poliovirus, a number of which encoded variant viral protein. Mammalian cells were cultured in vitro, where the cells included both parent poliovirus and variant poliovirus. A number of variant poliovirus that interfered with growth of the parent poliovirus in the cell were identified, and the mutations in the genomes of the variant poliovirus were identified. Mutations giving rise to the growth-interfering phenotype included mutations within the capsid coding region; mutations in the polymerase coding region; mutations in the protein primer 3B coding region and in the cis-acting replication element; and mutations in the 2A proteinase coding region. Because the variant proteins and/or RNAs interfered with growth of parent poliovirus, they were referred to as “dominant targets.” A dominant target would be a RNA or protein target that, when defective, dominantly interferes with the growth of non-defective virus.

It was also observed that the presence of a drug-sensitive virus inhibits growth of a drug-resistant virus in a cell in the presence of the drug, where the drug target is a dominant viral protein. FIG. 8 provides a schematic depiction of an exemplary assay for the effect of growth of a parent, drug-sensitive virus on growth of a variant, drug-resistant virus.

The present invention provides methods for identifying a candidate anti-viral agent, where the emergence of drug-resistant virus in an individual following administration of such an agent is reduced or delayed. A delay in emergence of drug-resistant virus allows an individual to mount an effective immune response to the virus. Thus, the individual can clear the infection before emergence of drug-resistant virus.

Methods of Identifying Candidate Anti-Viral Agents

The present invention provides methods of identifying a candidate anti-viral agent. The methods generally involve:

a) culturing a mammalian cell in vitro, where the mammalian cell comprises: i) a parent RNA virus, where growth of the parent RNA virus is inhibited by a test agent; and ii) a variant of the parent RNA virus, where growth of the variant RNA virus is resistant to the test agent; and

b) determining the effect, if any, of parent virus growth on growth of the variant virus during at least one replicative cycle.

If parent virus growth interferes with the variant virus growth during at least one replicative cycle, the test agent is considered a candidate anti-viral agent. In some embodiments, the parent virus growth interferes with growth of the variant virus during one replicative cycle in vitro or in vivo. In other embodiments, the parent virus growth interferes with growth of the variant virus during two, three, four, five, six, seven, eight, nine, ten, or more, replicative cycles in vitro or in vivo. Mammalian cells cultured in vitro and comprising both parent RNA virus and variant RNA virus, e.g., step (a), can be cultured in the presence or absence of the test agent. In some embodiments, step (a) is carried out in the absence of test agent.

A replicative cycle varies from virus to virus; replicative cycles for various RNA viruses are well known to those skilled in the art. For example, a single replicative cycle for poliovirus is about 6 hours in a mammalian cell in in vitro cell culture; and a single replicative cycle for HCV is about 48 hours in a mammalian cell in in vitro cell culture.

Whether a parent virus interferes with growth of a variant virus during at least one replicative cycle is readily determined using known methods. For example, growth of the parent virus and growth of the variant virus is determined in in vitro culture. The parent virus and the variant virus are introduced into a mammalian cell in in vitro culture; and the effect, if any, of growth of the parent virus on growth of the variant virus is determined.

In some embodiments, the effect of growth of the parent RNA virus on growth of the variant virus is determined by introducing both parent RNA virus and variant RNA virus into mammalian cells in in vitro culture, where the parent RNA virus is introduced into mammalian cells at a multiplicity of infection (MOI; or the average number of virus genomes per cell) of from about 1 to about 100, e.g., the parent RNA virus is introduced into mammalian cells at an MOI of from about 1 to about 5, from about 5 to about 10, from about 10 to about 20, from about 20 to about 30, from about 30 to about 40, from about 40 to about 50, from about 50 to about 70, or from about 70 to about 100; and the variant RNA virus is introduced into the mammalian cells at an MOI of from about 0.01 to about 1, e.g., the variant RNA virus is introduced into the mammalian cells at an MOI of from about 0.01 to about 0.05, from about 0.05 to about 0.07, from about 0.07 to about 0.1, from about 0.1 to about 0.2, from about 0.2 to about 0.5, or from about 0.5 to about 1.0. In some embodiments, the ratio of MOI of the parent virus to the variant virus is from about 5 to about 10⁴, e.g., from about 5 to about 10, from about 10 to about 10², from about 10² to about 5×10², from about 5×10² to about 10³, from about 10³ to about 5×10³, or from about 5×10³ to about 10⁴.

Growth of the drug-sensitive, parent RNA virus inhibits growth of the drug-resistant, variant RNA virus, e.g., the yield of drug-resistant, variant RNA virus is reduced. For example, growth of the drug-sensitive, parent RNA virus reduces the yield of drug-resistant, variant RNA virus per cell in in vitro mammalian culture by at least about 10%, at least about 20%, at least about 25%, at least about 30%, at least about 35%, at least about 40%, at least about 50%, at least about 60%, at least about 70%, at least about 80%, or at least about 90%, or more (e.g., at least about 95%, at least about 98%), during at least one replicative cycle (e.g., during one, two, three, four, five, or more replicative cycles). In some embodiments, yield of the drug-resistant, variant RNA virus is undetectable during one or more replicative cycles.

In some embodiments, step (a) is carried out in the absence of the test agent; and determining the effect of growth of the parent RNA virus on the variant virus is carried out using a plaque assay in the presence and in the absence of the test agent. In the absence of the test agent, the plaque assay provides the total number of viruses (parent+variant); and in the presence of the test agent, the plaque assay provides the number of variant virus. A control can be carried out, in which mammalian cells are culture in vitro, which mammalian cell comprise only the variant RNA virus; and the number of variant virus determined using a plaque assay in the presence and absence of the test agent. An example of such an assay is depicted schematically in FIG. 8.

Virus growth in vitro is readily measured using known assays. For example, in some embodiments, virus produced by a cell is harvested and virus stocks are determined by plaque assay. As another example, the parent virus and the variant virus could be constructed such that the parent virus includes a first reporter gene operably linked to a promoter; and the variant virus includes a second reporter gene operably linked to a promoter, where the first and second reporter genes encode products that are distinguishable from one another; and the effect of parent virus growth on variant virus growth is determined by detecting the first and second gene products. As another example, viral RNA is measured within the cell or in cell extracts or supernatants, virion production is measured by viral capsid or envelope protein in the supernatant.

Suitable reporter gene products include, but are not limited to, luciferase (e.g., firefly luciferase and derivatives thereof; Renilla luciferase and derivatives thereof); β-galactosidase; chloramphenicol acetyl transferase; glutathione S transferase; a green fluorescent protein (GFP), including, but not limited to, a GFP derived from Aequoria victoria or a derivative thereof, a number of which are commercially available; a GFP from a species such as Renilla reniformis, Renilla mulleri, or Ptilosarcus guernyi, as described in, e.g., WO 99/49019 and Peelle et al. (2001) J. Protein Chem. 20:507-519; any of a variety of fluorescent and colored proteins from Anthozoan species, as described in, e.g., Matz et al. (1999) Nature Biotechnol. 17:969-973, U.S. Patent Publication No. 2002/0197676, or U.S. Patent Publication No. 2005/0032085; a red fluorescent protein; a yellow fluorescent protein; a Lumio™ tag (e.g., a peptide of the sequence Cys-Cys-Xaa-Xaa-Cys-Cys, where Xaa is any amino acid other than cysteine, e.g., where Xaa-Xaa is Pro-Gly, which peptide is specifically bound by a fluorescein derivative having two As(III) substituents, e.g., 4′,5′-bis(1,3,2-dithioarsolan-2-yl)fluorescein; see, e.g., Griffin et al. (1998) Science 281:269; Griffin et al. (2000) Methods Enzymol. 327:565; and Adams et al. (2002) J. Am. Chem. Soc. 124:6063); and the like.

Growth of the drug-sensitive, parent RNA virus inhibits growth of the drug-resistant, variant RNA virus, e.g., the yield of drug-resistant, variant RNA virus is reduced. However, in some embodiments, production of variant viral RNA in the cytoplasm is unaffected by growth of the parent RNA virus. Whether production of variant virus RNA is reduced by growth of the parent RNA virus is readily determined using known assays. For example, RNA can be harvested from cells, and the amounts of parent and variant virus determined using a reverse-transcription/polymerase chain reaction (RT-PCR) method.

Suitable mammalian cells include primary cells and immortalized cell lines. Suitable mammalian cell lines include human cell lines, non-human primate cell lines, rodent (e.g., mouse, rat) cell lines, and the like. Suitable mammalian cell lines include, but are not limited to, HeLa cells (e.g., American Type Culture Collection (ATCC) No. CCL-2), CHO cells (e.g., ATCC Nos. CRL9618, CCL61, CRL9096), 293 cells (e.g., ATCC No. CRL-1573), Vero cells, NIH 3T3 cells (e.g., ATCC No. CRL-1658), Huh-7 cells, BHK cells (e.g., ATCC No. CCL10), PC12 cells (ATCC No. CRL1721), COS cells, COS-7 cells (ATCC No. CRL1651), RAT1 cells, mouse L cells (ATCC No. CCLI.3), human embryonic kidney (HEK) cells (ATCC No. CRL1573), HLHepG2 cells, and the like.

Parent RNA virus and variant RNA virus are introduced into the mammalian cell in vitro using any of a variety of methods, a number of which are well known in the art. For example, where the virus is an encapsidated virion, the virus can be introduced into a mammalian cell by infection. Where the virus is not encapsidated (e.g., the virus is in the form of non-encapsidated RNA or other non-encapsidated nucleic acid), the virus can be introduced into a mammalian cell by electroporation, transfection using DEAE-dextran, lipofection, and the like. DNA plasmids that encode the viral RNAs can also be used.

Virus growth in vivo is readily measured using known assays. For example, a mammal (e.g., a rodent) is infected with both parent and variant viruses, and virus growth is measured at various times post-infection. Virus may be harvested from specific tissues or organs (e.g. skin, brain, muscle, spleen, etc.) and the tissue disrupted (e.g. by sonication, freeze-thaw followed by mortar/pestle grinding, homogenization using blender or dounce, etc) and the emerging virus quantified by plaque assay or reporter gene assay as described above. Virus growth can be measured by plaque assay or by detection of viral products in the cell supernatant or by detection of intracellular viral genomes.

Parent RNA Virus

A parent RNA virus that is suitable for use in a subject method exhibits sensitivity to one or more test agents, e.g., a test agent inhibits growth of the parent RNA virus. Parent RNA viruses that are suitable for use in a subject method include wild-type RNA virus; wild-type RNA genome; any known serotype of an RNA virus; a DNA copy of a wild-type RNA genome; replication-competent variants of a wild-type RNA virus that retain sensitivity of a wild-type virus to a selected test agent and is capable of replication in a mammalian host cell; recombinant constructs comprising an RNA virus genome, or a DNA copy of an RNA virus genome; and sub-genomic replicons that retain sensitivity of a wild-type virus to a selected test agent.

Suitable parent RNA viruses include positive-strand RNA viruses and negative-strand RNA viruses. Suitable positive-strand RNA viruses include, but are not limited to, members of Picornaviridae, Flaviviridae, Togaviridae, Caliciviridae, Coronaviridae, and Retroviridae families. Positive-strand RNA viruses generally (except retroviruses) share the following features: replicate in the cytoplasm; genomic RNA serves as a message and is translated; genomic RNA is infectious; virions do not contain any enzymes; and viral proteins are translated as polyproteins.

Picornaviridae family members include, but are not limited to, members of genus Enterovirus (including poliovirus, enterovirus, coxsackievirus, echovirus); members of the genus Rhinovirus; members of the genus Hepatovirus (hepatitis A virus); encephalomyocarditis virus (EMCV); and foot-and-mouth disease virus (FMDV).

Flaviviridae family members include, but are not limited to, members of the genus flavivirus, e.g., Dengue virus, Yellow Fever Virus, St. Louis encephalitis virus, Japanese encephalitis virus, and West Nile virus; members of the genus hepacivirus, e.g., hepatitis C virus; and members of the genus pestivirus, e.g., bovine viral diarrhoea virus (BVDV).

Togaviridae family members include members of the genus alphavirus, e.g., Eastern encephalitis, western encephalitis, Sindbis, and Semliki forest viruses; and members of the genus rubivirus, e.g., rubella virus.

Retroviridae family members include members of the genus lentiviruses including, but not limited to, human immunodeficiency virus, simian immunodeficiency virus (SIV), and feline immunodeficiency virus.

Suitable negative-strand RNA viruses include, but are not limited to, Filoviridae family members (including Marburg virus, Ebola virus); Orthomyxoviridae family members (including influenza virus); Paramyxoviridae family members (including measles virus), and Rhabdoviridae family members (including rabies virus).

In some embodiments, a parent virus is a recombinant construct comprising an RNA virus genome, or a DNA copy of an RNA virus genome. The entire RNA genome (or a DNA copy thereof) may be present; however, the entire RNA genome (or a DNA copy thereof) need not be present in the recombinant construct. Suitable constructs include plasmid constructs that include a cDNA copy of an RNA virus genome, or a sub-genomic portion of an RNA virus genome. Sub-genomic replicons of HCV are known in the art and include, e.g., those described in U.S. Pat. No. 6,956,117.

Poliovirus is exemplified in the Examples, below. Nucleotide and amino acid sequences of poliovirus are known in the art. For example, the nucleotide sequence of poliovirus type 1 (Mahoney strain) is set forth in GenBank Accession No. NC_(—)002058; and is presented as SEQ ID NO:1; and amino acid sequences of the encoded proteins are set forth in GenBank Accession No. NP_(—)041277; and presented as SEQ ID NO:2. Amino acid sequences of individual proteins encoded by poliovirus type 1 (Mahoney) are provided in GenBank Accession Nos. NP_(—)740469 (VP2; SEQ ID NO:3; and FIG. 9); NP_(—)740470 (VP3; SEQ ID NO:4; and FIG. 10); NP_(—)740471 (VP1; SEQ ID NO:5; and FIG. 11); NP_(—)740477 (2A; SEQ ID NO:6; and FIG. 12); NP_(—)740472 (2B; SEQ ID NO:7; and FIG. 13); NP_(—)740473 (2C; SEQ ID NO:8; and FIG. 14); NP_(—)740474 (3A; SEQ ID NO:9; and FIG. 15); NP_(—)740475 (3B; SEQ ID NO:10; and FIG. 16); NP_(—)740476 (3C; SEQ ID NO:11; and FIG. 17); NP_(—)740478 (3D; SEQ ID NO:12; and FIG. 18); and NP_(—)740468 (VP4; SEQ ID NO:13 and FIG. 19). Sequences of poliovirus type 3 (Fox strain) are set forth in GenBank Accession No. AY359875. Sequences of poliovirus type 3 (Sabin) are set forth in GenBank Accession Nos. X00596 and P03302.

Variant RNA Virus

A variant RNA virus is a variant of any of the above-described parent RNA virus that, unlike the parent RNA virus, is resistant to a particular agent, e.g., the variant virus grows in a mammalian cell in the presence of a selected test agent. Thus, e.g., where growth of a parent RNA virus is inhibited by a selected test agent, growth of a variant of the parent RNA virus is resistant to the test agent.

A variant RNA virus genome comprises one or more changes in nucleotide sequence relative to the nucleotide sequence of the parent RNA virus genome. The one or more changes in nucleotide sequence result in a change in the RNA and/or an encoded protein that inhibits growth of the parent virus RNA in a mammalian cell containing both parent virus RNA and variant virus RNA when the cell is cultured in the absence of a test agent. Thus, the one or more changes in nucleotide sequence result in a change that provides for a dominant-negative phenotype when parent virus and a variant virus are present together in the same mammalian cell in the absence of any test agent. In some embodiments, the one or more changes in nucleotide sequence result in a change in viral RNA or viral-encoded protein such that the variant virus, is non-viable when grown in the absence of parent virus (or any other wild-type virus or any other virus that compensates for the mutation) in a mammalian cell.

As noted above, the one or more changes in nucleotide sequence result in a change that provides for a dominant-negative phenotype when parent virus and a variant virus are present together in the same mammalian cell in the absence of any test agent. Thus, e.g., when variant RNA virus and parent virus are introduced into mammalian cells in in vitro cell culture in the absence of test agent, and at an excess of variant virus over parent virus, the variant virus reduces parent viral growth by at least about 10%, at least about 20%, at least about 25%, at least about 30%, at least about 35%, at least about 40%, at least about 50%, at least about 60%, at least about 70%, at least about 80%, or at least about 90%, or more (e.g., at least about 95%, at least about 98%), during at least one replicative cycle.

In some embodiments, the variant RNA virus comprises one or more changes in nucleotide sequence, compared to the nucleotide sequence of the parent RNA virus, where the one or more nucleotide sequence changes alter the amino acid sequence of an encoded viral oligomeric protein, such that the virus produces a variant viral oligomeric protein in the cytoplasm of a mammalian cell. Variant viral oligomeric proteins include variants having one or more amino acid sequence changes relative to the parent oligomeric protein, where amino acid sequence changes include insertions, deletions, truncations, substitutions, etc. In some embodiments, a variant viral oligomeric protein has a single amino acid substitution, compared with the parent viral oligomeric protein. The variant oligomeric protein inhibits growth of a parent virus present in the same cell. Oligomeric viral proteins that, when mutated, exhibit a dominant negative phenotype, include, but are not limited to, viral capsid proteins; membrane-associated proteins; and polymerases (e.g., RNA-dependent RNA polymerase).

In some embodiments, the variant RNA virus comprises one or more changes in nucleotide sequence, compared to the nucleotide sequence of the parent RNA virus, where the one or more nucleotide sequence changes alter the amino acid sequence of an encoded viral trans-acting protein. Variant viral trans-acting proteins include variants having one or more amino acid sequence changes relative to the parent trans-acting protein, where amino acid sequence changes include insertions, deletions, truncations, substitutions, etc. In some embodiments, a variant viral trans-acting protein has a single amino acid substitution, compared with the parent viral trans-acting protein. The variant trans-acting protein inhibits growth of a parent virus present in the same cell. Trans-acting proteins include, but are not limited to, protein primer; polymerases, proteinases that cleave polyproteins; RNA helicases; membrane associated proteins, and the like.

Identification of Dominant Negative Protein Targets

Suitable viral variants can be identified using a method as follows. In some embodiments, a target for anti-viral drug intervention is identified by a method comprising: a) introducing a mutation into a coding region of the genome of a parent RNA virus, generating a variant RNA virus comprising a variant coding region; b) co-transfecting a mammalian cell in vitro with the parent RNA virus and the variant RNA virus; and c) determining the effect, if any, of the variant RNA virus on growth of the wild-type RNA virus. Inhibition of growth of the parent RNA virus by the variant RNA virus indicates that the product of the variant coding region is a suitable target for anti-viral drug intervention.

Methods for introducing mutations into a coding region of a viral genome are well known in the art, and any known method can be used. Mutations that are introduced into the viral genome result in one or more changes in amino acid sequence of an encoded viral protein, relative to the amino acid sequence of the viral protein encoded by the parent RNA virus. Mutations include random mutations and directed (non-random) mutations.

Methods of mutating a nucleic acid are well known in the art and include well-established chemical mutation methods, radiation-induced mutagenesis, and methods of mutating a nucleic acid during synthesis. Chemical methods of mutating DNA include exposure of DNA to a chemical mutagen, e.g., ethyl methanesulfonate (EMS), methyl methanesulfonate (MMS), N-nitrosourea (ENU), N-methyl-N-nitro-N′-nitrosoguanidine, 4-nitroquinoline N-oxide, diethylsulfate, benzopyrene, cyclophosphamide, bleomycin, triethylmelamine, acrylamide monomer, nitrogen mustard, vincristine, diepoxyalkanes (e.g., diepoxybutane), ICR-170, formaldehyde, procarbazine hydrochloride, ethylene oxide, dimethylnitrosamine, 7,12 dimethylbenz(a)anthracene, chlorambucil, hexamethylphosphoramide, bisulfan, and the like. Radiation mutation-inducing agents include ultraviolet radiation, γ-irradiation, X-rays, and fast neutron bombardment. Mutations can also be introduced into a nucleic acid using, e.g., trimethylpsoralen with ultraviolet light. Random or targeted insertion of a mobile DNA element, e.g., a transposable element, is another suitable method for generating mutations. Mutations can be introduced into a nucleic acid during amplification in a cell-free in vitro system, e.g., using a polymerase chain reaction (PCR) technique such as error-prone PCR. Non-random mutations can be introduced using a PCR method, where a primer is used that has one or more nucleotide differences from the template. Mutations can be introduced into a nucleic acid in vitro using DNA shuffling techniques (e.g., exon shuffling, domain swapping, and the like). Mutations can also be introduced into a nucleic acid as a result of a deficiency in a DNA repair enzyme in a cell, e.g., the presence in a cell of a mutant gene encoding a mutant DNA repair enzyme is expected to generate a high frequency of mutations (i.e., about 1 mutation/100 genes-1 mutation/10,000 genes) in the genome of the cell. Examples of genes encoding DNA repair enzymes include but are not limited to Mut H, Mut S, Mut L, and Mut U, and the homologs thereof in other species (e.g., MSH 1-6, PMS 1-2, MLH 1, GTBP, ERCC-1, and the like). Methods of mutating nucleic acids are well known in the art, and any known method is suitable for use. See, e.g., Stemple (2004) Nature 5:1-6; Chiang et al. (1993) PCR Methods Appl 2(3): 210-217; Stemmer (1994) Proc. Natl. Acad. Sci. USA 91:10747-51; and U.S. Pat. Nos. 6,033,861, and 6,773,900.

The variant viral-encoded protein will have one or more changes in amino acid sequence, compared to the amino acid sequence of the same protein encoded by the parent RNA virus. Variants will have one or more amino acid substitutions, insertions, or deletions compared to the amino acid sequence of the same protein encoded by the parent RNA virus. In some embodiments, the variant viral-encoded protein will have a single amino acid substitution, compared to the amino acid sequence of the same protein encoded by the parent RNA virus. Variant RNA sequences will have one or more nucleotide changes relative to parent RNA virus.

Test Agents

The terms “candidate agent,” “test agent,” “agent,” “substance,” and “compound” are used interchangeably herein. Test agents encompass numerous chemical classes, typically synthetic, semi-synthetic, or naturally-occurring inorganic or organic molecules. Candidate agents include those found in large libraries of synthetic or natural compounds. For example, synthetic compound libraries are commercially available from Maybridge Chemical Co. (Trevillet, Cornwall, UK), ComGenex (South San Francisco, Calif.), and MicroSource (New Milford, Conn.). A rare chemical library is available from Aldrich (Milwaukee, Wis.). Alternatively, libraries of natural compounds in the form of bacterial, fungal, plant and animal extracts are available from Pan Labs (Bothell, Wash.) or are readily producible. The term “test agent” excludes WIN-51711.

Candidate agents may be small organic or inorganic compounds having a molecular weight of more than 50 and less than about 2,500 daltons. Candidate agents may comprise functional groups necessary for structural interaction with proteins, particularly hydrogen bonding, and may include at least an amine, carbonyl, hydroxyl or carboxyl group, and may contain at least two of the functional chemical groups. The candidate agents may comprise cyclical carbon or heterocyclic structures and/or aromatic or polyaromatic structures substituted with one or more of the above functional groups. Candidate agents are also found among biomolecules including peptides, saccharides, fatty acids, steroids, purines, pyrimidines, derivatives, structural analogs or combinations thereof. In some embodiments, test agents include neutralizing antibodies. In some embodiments, neutralizing antibodies are specifically excluded.

Selecting for Drug-Resistant Viral Variants

As noted above, a subject method for identifying a candidate anti-viral agent involves culturing a mammalian cell in vitro, where the mammalian cell comprises a drug-sensitive parent RNA virus (e.g., a parent RNA virus that exhibits growth inhibition in the presence of a test agent) and a drug-resistant variant RNA virus (e.g., a variant virus, where growth of the variant virus is resistant to the test agent). Selection of a variant RNA virus that is resistant to a test agent can be carried out using standard methods. For example, mammalian cells comprising a parent RNA virus are cultured in vitro in the presence of a test agent that inhibits growth of the parent RNA virus. Variants are selected that are not growth-inhibited, and therefore are resistant to the test agent.

EXAMPLES

The following examples are put forth so as to provide those of ordinary skill in the art with a complete disclosure and description of how to make and use the present invention, and are not intended to limit the scope of what the inventors regard as their invention nor are they intended to represent that the experiments below are all or the only experiments performed. Efforts have been made to ensure accuracy with respect to numbers used (e.g. amounts, temperature, etc.) but some experimental errors and deviations should be accounted for. Unless indicated otherwise, parts are parts by weight, molecular weight is weight average molecular weight, temperature is in degrees Celsius, and pressure is at or near atmospheric. Standard abbreviations may be used, e.g., bp, base pair(s); kb, kilobase(s); pl, picoliter(s); s or sec, second(s); min, minute(s); h or hr, hour(s); aa, amino acid(s); kb, kilobase(s); bp, base pair(s); nt, nucleotide(s); i.m., intramuscular(ly); i.p., intraperitoneal(ly); s.c., subcutaneous(ly); and the like.

Example 1 Identification of Dominant Targets

Methods

Design and Characterization of Mutant Genomes Used for Dominant Inhibitor Screen.

To amass a collection of nonviable mutant poliovirus genomes, previously characterized lethal mutations, or mutations that were designed to destabilize the structure of the encoded viral protein product, were introduced using two strategies. The first strategy was to disrupt the hydrophobic core of the encoded protein by reducing the size of an amino acid side chain predicted to be inaccessible to solvent or by changing it to Pro. A second strategy was to disrupt predicted α-helices by altering Leu and Ser residues predicted to reside in α-helices to Pro. A computer algorithm, PredictProtein³³, was used to predict the likely solvent accessibility and α-helicity of each amino acid position within the poliovirus type 1 (Mahoney) polyprotein in its native, folded state. Single amino acids presumed to be within the hydrophobic protein core by virtue of displaying a PredictProtein score greater than five were altered by introducing a single U-to-C transition mutation into the viral genomes, using PCR-mediated site-directed mutagenesis³⁴. The PredictProtein method was used instead of known three-dimensional structures to simulate the knowledge base of other, less well-characterized, positive-strand RNA viruses.

The viability of each mutant genome was tested by transfecting 60 mm plates of subconfluent S3 HeLa monolayers with 1 μg of in vitro transcribed mutant RNA using DEAE-dextran (see below). Viral stocks were harvested after a single replicative cycle (10 hrs at 32.5° C.) by pelleting the cells at 200×g, washing and lysing by freeze-thaw in 1 ml of PBS+ (phosphate buffered saline with 0.1% CaCl₂ and MgCl₂), and lysing the cells by freeze-thaw treatment. Stocks were then titered as previously described³⁵. Mutant genomes that produced no detectable plaques in this assay were defined as nonviable; 1 μg of wild-type RNA typically yielded between 4×10⁵-1×10⁶ PFU under these conditions. The results of the engineered and previously characterized mutations are described in Tables 1 and 4. Reversion of mutant genomes to produce wild-type virus was not detected using DEAE-dextran transfections.

RNA Transcription and Co-Transfections

Plasmids containing the poliovirus 1 genome (pGEM-PV1) under the control of a T7 promoter were linearized by Eco R1 restriction digestion (New England Biolabs, Beverly, Mass.), purified by agarose gel electrophoresis, and used as a template for transcription using the Ribomax (Promega, Madison, Wis.) T7 transcription kit according to the manufacturer's standard protocol. A control RNA, R2-PvuII (see below), was made from pGEM-PV1 cDNA that lacks capsid encoding nucleotides 1175 to 2956 and linearized with PvuII, which cleaves within the coding region of 3D polymerase. All transcription reactions were extracted twice with acid phenol-chloroform-(5:1, pH 4.5, Ambion) to remove protein and template DNA, and precipitated with a half volume of 7.5 M ammonium acetate and 2.5 volumes ethanol. Precipitated pellets of RNA were then resuspended in water and applied to a P30 size exclusion column (Biorad, Hercules, Calif.) to remove unincorporated nucleotides and the eluate collected according to the manufacturer's protocol. The integrity of transcribed RNA was verified by denaturing formaldehyde agarose gel electrophoresis and the concentration determined by measuring absorbance at 260 nm. The preparations were reprecipitated, aliquoted and stored at −80° C. Transcription reactions were also performed in the presence of α³²P-UTP to verify that RNA amounts measured by O.D. at 260 nm represented RNA transcripts and not unincorporated nucleotides.

The effect of each mutant genome on the growth of wild-type virus was tested by co-transfecting 60 mm plates of subconfluent S3 HeLa monolayers with 1 μg of in vitro transcribed mutant RNA and 100 ng of wild-type poliovirus RNA using DEAE-dextran (average molecular weight=500,000; Sigma, St. Louis, Mo.) as described previously⁸. After 10 hours incubation of each transfection at 32.5° C., cells were harvested, virus stocks produced, and plaque assays performed as described³⁵.

In Vitro Translation of Poliovirus RNAs and α-VP1 Immunoprecipitation of Translation Reactions

Poliovirus RNAs were transcribed as for transfections. HeLa S10 extracts were prepared as previously described³⁶, and a 25 μl reaction was programmed with 5 μg of RNA, 50% HeLa extract, 1 μl ³⁵S-Met Express label (NEN), 0.5 μl RNasin (Promega), and 0.5 μl 1 mM of each amino acid except methionine. Each reaction was incubated for 2 hours at 30° C., and then stopped by addition of 2×-lysis buffer (2% Triton X-100 prepared in TBS: 10 mM Tris-Cl pH 8.0, 140 mM NaCl, and 0.025% NaN₃). Each reaction was then centrifuged at 10,000×g for 30 minutes at 4° C. To clear the supernatant, 1/20^(th) volume of a 50% slurry of protein-G sepharose beads (Gibco-BRL, Grand Island, N.Y.) was added to the supernatant, incubated at room temperature for 2 hrs, and pelleted by 1 min. centrifugation at 200×g. The supernatant was transferred to a new tube precoated with lysis buffer that contained 200 μl of dilution buffer (0.1% Triton X-100 prepared in TBS). Monoclonal mouse α-VP1 antibody (3 μg; Chemicon, Temecula, Calif.) was added to each reaction and incubated 1.5 hours at 4° C. The beads were pelleted at 200×g and washed twice with dilution buffer, once with TBS, and once with 50 mM Tris-Cl (pH 6.8). Proteins in the samples were then separated on a 10% polyacrylamide gel by SDS-PAGE analysis.

Super-Infections

Temperature-sensitive viruses were either previously described²⁴ or generated during the screening of nonviable poliovirus type 1 (Mahoney) mutants (Table 3). HeLa cell monolayers were infected at an MOI=100 PFU/cell with each temperature-sensitive virus at a semi-permissive temperature of 37° C. in PBS+. After virus adsorption (30 minutes), serum supplemented DMEM was added with 2 mM guanidine hydrochloride to block viral RNA synthesis. At two hours post-infection, the media was removed and cells were super-infected with wild-type virus at an MOI of 0.5 PFU/cell. Incubation at 37° C. was continued for 4 hours in the absence of guanidine, whereupon cells were harvested and virus stocks were made as described above. Virus titers were determined at 39° C.

Co-Infections of WIN-Sensitive and WIN-Resistant Viruses

WIN-resistant Sabin 3 virus was engineered by introducing a mutation, VP1-I192F, into a cDNA encoding the attenuated Sabin type 3 poliovirus. Infections and co-infections of HeLa cell monolayers at the indicated MOI's for 30 minutes at 37° C. Serum-supplemented DMEM (10%) with or without 2 μg/ml final concentration WIN-51711 was added to the cells, and incubation continued at 37° C. for 6 hours. Cells were then harvested and virus stocks titered by plaque assay in the absence and presence of 2 μg/ml WIN-51711 in the agar overlay.

RT-PCR of WIN-R and Wild-Type RNA

Co-infections and single infections of wild-type and WIN-resistant viruses were performed as described above in the absence of WIN-51711. At 5 hours post-infection, cells were harvested and processed by the addition of 1 ml Trizol (Invitrogen, Carlsbad, Calif.) to each plate and incubation for 5 min. at room temperature. The solution was extracted with 0.2 ml chloroform and the supernatants were collected. At this time, “mix” samples were created by combining equal volumes of supernatants derived from single infections (see FIGS. 6 c,d) Nucleic acids were collected by the addition of isopropanol (70%), pelleting by centrifugation, washing with 70% ethanol, repelleting, and resuspension in 50 μl 10 mM HEPES-KOH (pH 7.5).

Each reverse transcriptase reaction contained 5 μl RNA sample in a final volume of 10 μl using AMV-RT High Concentration (Promega, Madison, Wis.) as recommended by the manufacturer. PCR reactions (50 μl total volume) were composed of 5 μl of a reverse transcriptase reaction and performed as previously described³⁷. Reactions were cycled 35 times (94° C., 1 minute; 54° C., 1 minute; 72° C. 1 minute). Primers used amplified a region from nucleotide 2967 to 3241 of the type 3 genome. Two TfiI restriction sites (at nucleotides 3048 and 3149) exist within the wild-type PCR product, but the 3048 site is disrupted by the VP1 I192F WIN-R mutation. PCR reactions were brought to 200 μl volumes and digested with 25 units TfiI for two hours at 65° C. Reactions were then ethanol precipitated and analyzed by PAGE on a 5% polyacrylamide, 8M urea gel.

Results

Design and Characterization of Nonviable Poliovirus Mutations

To search for dominant alleles in a comprehensive, genome-wide manner, a battery of lethally mutated genomes spanning the poliovirus coding region was constructed (Table 1). The construction of each mutant genome was guided either by a previously described mutation or by a strategy to disrupt the structure of the encoded protein. By targeting predicted hydrophobic cores or α-helices (see Methods), 24 individual U-to-C mutations were introduced into an infectious poliovirus cDNA and the viability of each mutant viral genome was tested. The results of these transfections and the rationale for each mutation are shown in Table 1. TABLE 1 Mutant poliovirus genomes constructed for use in the dominance screen. Codon Viability Mutant Change Rationale (PFU/ml) Previously characterized mutant poliovirus genomes VP2-S1P UCG→CCG Maturation cleavage (Ansardi <5 and Morrow 1995) VP2-S243P UCC→CCC Reynolds et al. 1991 <5 2A-C109R UGU→CGU Catalytic proteinase cysteine <5 (Yu and Lloyd 1991) 3B-Y3H UAC→CAC Uridylylation site (Rothberg <5 et al. 1978; Ambros and Baltimore 1978) 3C-C147R UGU→CGU Catalytic proteinase cysteine <5 (Hammerle et al. 1991) 3D-F30S UUC→UCC Fingers-thumb interaction <5 (Hobson et al. 2001; Hansen et al. 1997) 3D-S291P UCA→CCA Burns et al. 1989 <5 CRE-C4465U/ None Goodfellow et al. 2000 <5 U4483C* CRE-G4462A/ None Goodfellow et al. 2000 <5 U4483C* Designed mutant poliovirus genomes VP2-F260S UUC→UCC Hydrophobic <5 VP3-F118S UUU→UCU Hydrophobic <5 VP3-L211S CUU→CCU Hydrophobic <5 VP1-L118P UUA→UCA Hydrophobic & Helix <5 2A-S74P UCC→CCC Helix t.s. 2A-L98P CUC→CCC Hydrophobic <5 2A-F133S UUU→UCU Hydrophobic <5 2B-F13S UUU→UCU Hydrophobic & Helix <5 2B-F17S UUU→UCU Hydrophobic & Helix <5 2C-F28S UUC→UCC Hydrophobic & Helix t.s. 2C-L93P CUU→CCU Hydrophobic <5 2C-F242S UUU→UCU Hydrophobic <5 2C-F328S UUU→UCU Hydrophobic & Helix <5 3A-L8S UUG→UCG Hydrophobic <5 3A-L24S UUG→UCG Hydrophobic & Helix <5 3A-F83S UUU→UCU Hydrophobic <5 3C-L70P CUU→CCU Hydrophobic <5 3C-L102S UUG→UCG Hydrophobic <5 3D-F34L UUU→CUU Hydrophobic t.s. 3D-L107P CUA→CCA Hydrophobic <5 3D-F191S UUU→UCU Hydrophobic & Helix <5 3D-F246S UUC→UCC Hydrophobic & Helix <5 3D-F296S UUU→UCU Hydrophobic & Helix <5 3D-Y326H UAU→CAU Hydrophobic <5 *Mutants in the CRE (cis-acting replication element) are double mutants because viruses containing either single mutation were viable. CRE mutations are non-coding mutations in the 2C coding region.

For 21 of the 24 designed mutations, no virus was detected upon a single cycle of growth after RNA transfection, indicating an absence of reversion to wild-type virus under these transfection conditions. The three remaining mutations gave rise to viable viruses with temperature-sensitive phenotypes that were characterized further (Table 3). For both previously published and designed mutations, only mutant genomes that displayed a 100,000-fold or greater reduction in plaque formation after RNA transfection were determined to be nonviable and used in the screen for dominant negative poliovirus alleles, which identified four classes of strongly dominant alleles.

Dominant-Negative Alleles in Capsid Coding Regions

To test the ability of nonviable mutant genomes to affect wild-type viral growth, nonviable and wild-type viral RNAs were co-transfected into HeLa cells, the intracellular virus was harvested after a single replicative cycle, and the resulting wild-type virus stocks were titered. To both mimic a scenario in which a drug-resistant genome emerges from a drug-sensitive population, and to optimize co-transfection conditions, a ten-fold excess of the nonviable genome was chosen. Total yeast tRNA was substituted for mutant RNA in the positive wild-type control. Under these conditions, a transfection of approximately 2×10⁶ cells with 100 ng wild-type RNA typically yielded a virus stock of 50,000-200,000 PFU (plaque-forming units)/ml.

The effect of a known inhibitor of poliovirus RNA replication, an RNA transcript (R2-PvuII) made in vitro from a poliovirus cDNA template cleaved with Pvu II⁷, was tested to ensure that the transfection protocol used led to co-transfection of the wild-type and potentially inhibitory genomes. When co-transfected with wild-type viral RNA, a ten-fold excess of R2-PvuII RNA inhibited wild-type growth (FIG. 1 b), as reported previously⁸. Although the mechanism by which R2-PvuII RNA inhibits the growth of wild-type RNA is not known, the greater than 20-fold inhibition of wild-type growth observed argues that at least 95% of the cells that contained wild-type viral RNA also contained the co-transfected inhibitor RNA.

Co-transfection of several of the lethally mutated RNAs, for example fs-2956 and 3A-L24S, either had little effect or caused a slight increase in wild-type yield (FIG. 1 b). Although 3A-L24S genomes contained a lethal point mutation in one coding region, it is likely that other functional trans-acting proteins produced from these mutant genomes provide helper functions for the wild-type genomes. The frame-shift control, fs-2956, occurs in the center of the VP1 coding region, and produces a truncated wild-type capsid region with termination of translation at a stop codon at nucleotide 3129. This construct appears to have also provided a helper function by the production of capsid proteins.

In contrast, all four genomes that contained lethal mutations within the capsid coding region, VP2-S1P, VP2-S243P, VP3 L211S, and VP1-L118P, reduced wild-type viral growth approximately 10- to 20-fold (FIGS. 1 a,c), which was the same extent of inhibition observed for the R2-PvuII co-transfection control. On average, capsid mutant genomes inhibited wild-type growth to 7% of wild-type growth alone.

FIGS. 1 a-d. Dominant inhibitor screen for capsid-coding genome regions. (a) Schematic of mutant genomes tested as dominant inhibitors of wild-type virus growth. (b) Validation of dominant inhibitor screen for R2-PvuII, a known RNA inhibitor of poliovirus growth, and two mutant alleles, a frameshift mutation at nucleotide 2956 (fs-2956) and 3A-L24S, that each provide apparent helper function. The average of each set of replicate experiments (with standard error) is shown below each set of replicates normalized to the average of the wild-type poliovirus RNA with carrier tRNA control. (c) Mutant capsid alleles mapped to the crystal structure of capsid proteins (VP4 in cyan, VP1 in yellow, VP2 in magenta, VP3 in salmon)⁴⁸. (d) The effect of co-transfecting the indicated RNAs with wild-type RNA on yield of wild-type poliovirus is shown as in b.

Allele-Specific Inhibition by Mutations in the 3D Polymerase Coding Region

Given the known ability of the poliovirus RNA-dependent RNA polymerase to oligomerize¹⁰, the dominance of five different non-viable alleles that contained mutations in the polymerase coding region was tested (FIGS. 2 a,b). One allele, S291P, diminished wild-type viral growth to 1% of the control (FIG. 2 c), and thus exerted a larger dominant effect than the R2-PvuII negative control. Two other alleles, F30S and F191S, diminished wild-type growth to 29% and 13%, respectively, and thus were co-dominant. Other alleles showed variable intermediate or helper effects and were deemed recessive. Mapped onto the fully resolved 3D polymerase structure¹¹, the majority of mutated residues cluster in the hydrophobic core of the fingers domain, while 3D-F30S is located at the interface between the “finger” and “thumb” domains (FIG. 2 b). While verifying that the screen adequately identifies residues involved in hydrophobic interactions, the variability in observed dominance of mutant 3D polymerase alleles may reflect varying degrees of protein stability or the oligomerization potential of 3D polymerase or its precursors. Alternatively, the allele-specificity of these alleles may also reflect their mixed effects on either 3D polymerase or its precursor, 3CD protease.

FIGS. 2 a-c. Effect of mutations in 3D polymerase on the yield of wild-type virus during co-transfection. (a) Schematic diagram of poliovirus genomes indicate locations of coding regions for mutant 3D polymerase alleles. (b) Mutant alleles mapped to the three-dimensional structure of 3D polymerase¹¹. (c) Parallel co-transfection experiments with wild-type RNA and viral genomes containing several different mutations in the coding region for 3D polymerase are shown as in FIGS. 1 a-d.

Dominant Mutations of the Protein Primer 3B and Cis-Acting Replication Element (CRE)

A surprising result came from the trans-dominant effects of mutations in the CRE, the nominally cis-acting RNA sequence that templates VPg uridylylation in vitro¹⁴, and 3B (the VPg coding region), shown in FIG. 3 a. Two non-coding double mutations in the CRE, G19A/U40C and C22U/U40C, as well as a mutation of the genome-linked structural protein, 3B-Y3H (FIG. 3 b), strongly inhibited growth of co-infecting wild-type virus. The degree of inhibition was similar to, or greater than, that exerted by R2-PvuII, the co-transfection control, or any of the capsid alleles.

A “classic” dominant negative allele is one in which the function of a protein or sequence element is disrupted while an associative property, such as a protein-protein or protein RNA interaction, is retained¹⁷. For the dominant negative alleles of the CRE, the stem-loop structure is predicted to be maintained (FIG. 3 a)¹⁶. The introduction of eight non-coding mutations into the CRE, however, is predicted to completely disrupt stem-loop structure and presumably any structure-specific associations¹⁸. This predicted loss-of-function CRE allele (“l.o.f. CRE”) did not inhibit wild-type growth (FIG. 3 c). Therefore, dominant inhibition by mutant CRE-containing genomes is allele-specific, presumably requiring an intact RNA stem-loop structure to form an inhibitory complex. Presumably such a complex would involve 3CD, a precursor known to bind RNA sequences in the 5′ UTR as well as the CRE.^(15,16).

FIGS. 3 a-c. Dominant inhibitor screen in the CRE and 3B-coding regions. (a) A schematic diagram of the predicted secondary structure of the wild-type CRE, which resides in the coding region of 2C, with indicated G19A, C22U, and U40C mutations used in the dominance screen. CRE mutants G19A/U40C and C22U/U40C correspond to genomic nucleotide positions 4462/4483 and 4465/4483, respectively, and are previously published non-coding mutations⁴⁷. 3B encodes the protein primer, VPg, to which uridyl residues are attached at Tyr-3. Asterisks (*) denote multiple non-coding nucleotides mutated to form the “l.o.f.”, or putative “loss-of-function” CRE used in c¹⁸. (b) Co-transfection experiments were performed mixing wild-type and mutant genomes that contain the indicated mutant alleles. (c) Specificity of mutant CRE alleles. Co-transfections of wild-type and mutant genomes that contain the specified CRE alleles were performed as in FIGS. 1 a-d. Mutations that specify the “l.o.f.” mutant genome is illustrated in a.

Allele-Specific Dominance in 2A Proteinase Coding Region

Two different mutations tested in the 2A proteinase coding region (FIGS. 4 a,b) showed pronounced dominance (2A-L98P and 2A-C109R) when compared to fs-2956 (FIG. 4 c). The dominant phenotype correlated with protease deficiency: FIG. 4 d shows the accumulation of uncleaved VP1-2A precursor for the 2A-L98P and 2A-C109R mutations, whereas wild-type and genomes containing mutations in the capsid coding region did not^(19,20). Further experiments using only the VP1-2A region expressed in vitro recapitulated this protease-deficient phenotype.

The dominance of protease-deficient 2A alleles was surprising at first, because 2A proteinase is known to be a monomeric enzyme with several viral and cellular substrates. However, its activity at the VP1-2A cleavage site is thought to be obligately intramolecular, because cleavage is unaffected by α-2A antibodies, and occurs more rapidly and with more specificity than intermolecular 2A proteinase cleavage^(21,22). A 2A protein that lacked enzymatic activity could therefore yield an uncleaved VP1-2A fusion molecule that functioned as a dominant negative inhibitor of wild-type growth, like a mutant capsid protein. To test this hypothesis explicitly, the VP1-2A cleavage site was mutagenized to allow accumulation of the uncleaved VP1-2A product from mutant genomes, and studied its effect on the growth of co-infecting wild-type virus. Two introduced mutations reported to abrogate 2A-mediated cleavage of the VP1-2A cleavage site, VP1-Y302P and VP1-T301R²², were also dominant (FIG. 4 e). Therefore, uncleaved VP1-2A is toxic to co-infecting wild-type virus, and its accumulation in cells infected with 2A proteinase-deficient mutant viruses is a likely mechanism for its genetic dominance.

FIGS. 4A-E. Dominant inhibitor screen for 2A proteinase and VP1-2A cleavage site mutant alleles. (a) Schematic diagram of poliovirus genomes indicate relative locations of coding regions for 2A and VP1, and the 2A proteinase and VP1-2A cleavage site mutants used in the dominant inhibitor screen. (b) 2A proteinase amino acid residues targeted to generate nonviable mutations used in the dominance inhibitor screen and superinfection assay are mapped onto the crystal structure of 2A proteinase from human rhinovirus 2, a closely related homolog of poliovirus 2A proteinase⁴⁹. (c) Co-transfections were performed as in FIG. 1 and the resulting wild-type virus yields of replicate experiments are shown. Normalization of these values as a percentage of the wild-type RNA with tRNA carrier control is shown at bottom with standard error. (d) HeLa cytoplasmic lysates were programmed with the indicated poliovirus RNAs and labeled using ³⁵S-methionine (Methods), immunoprecipitated with a monoclonal anti-VP1 antibody, and separated on a 10% SDS-PAGE gel. VP1 and VP1-2A mobilities are marked. The asterisk (*) denotes a higher molecular weight species abundant in 2A-L98P and 2A-C109R reactions. (e) Testing for dominance of genomes with mutated VP1-2A sites was performed as in FIGS. 1 a-d.

Translation and RNA Replication Requirements for Dominance

Genomes that contain mutant capsid alleles are usually known to be competent for RNA replication; therefore, the observed dominance of genomes with capsid mutations may be augmented by the replication of the mutant genomes, resulting in the production of high concentrations of mutant capsid proteins. To determine whether or not mutant capsid alleles require RNA replication to exert their dominant effect, a second mutation, ΔGUA₃, was introduced into one of the dominant mutant genomes. The deletion of nucleotides 7418-7422 in the 3′-non-coding region (FIG. 5 a) is known to severely diminish negative-strand RNA synthesis⁹. As shown in FIG. 5 b, the doubly mutant genome VP2-S243P/ΔGUA₃ did not inhibit wild-type virus growth. Therefore, the dominance of the VP2-S243P allele, and probably the other capsid alleles as well, requires replication of the mutant RNA genome, presumably leading to increased accumulation of the VP2-S243P mutant capsid proteins.

As with capsid alleles, 3D polymerase functions can be rescued in trans by polymerase molecules encoded by the co-transfected wild-type genomes, thus enabling genomes harboring mutant 3D polymerase alleles to replicate^(8,12,13). Whether the dominance of the 3D-S291P allele required RNA replication of its genome by introducing the ΔGUA₃ deletion was tested. As shown in FIG. 5 c, the 3D-S291P allele was no longer dominant when RNA replication of its genome was inhibited. Like the inhibitory effects of mutant capsid proteins, the high concentration of defective polymerases provided by a replicating genome is needed to inhibit the growth of co-infecting wild-type virus.

To test whether the observed dominance of the 3B mutation required RNA replication of the nonviable genome to exert dominance, the ΔGUA₃ mutation was introduced. Like dominant capsid and polymerase alleles, the 3B-Y3H allele required replication of its RNA genome to exhibit dominant negative effects on wild-type growth (FIG. 5 d).

However, when the same experiment was performed with dominant negative CRE allele C22U/U40C, the triple mutant C22U/U40C/ΔGUA₃ was still inhibitory, arguing that the mutant CRE structure was toxic at lower concentrations than the defective capsid, polymerase, or VPg proteins (FIG. 5 e). That such a dominant negative effect could occur without RNA replication is not without precedent since our dominant negative control for co-transfection, R2-PvuII, lacks a 3′-non-coding region and also presumably lacks the ability to replicate. To determine whether the CRE RNA alone was the inhibitory moiety, it was undertaken to determine whether a non-translatable CRE-C22U/U40C genome was dominant. To this end, the initial methionine of the poliovirus polyprotein was mutated to an amber stop codon (VP4-M1stop), and introduced into a genome containing the C22U/U40C CRE allele. As shown in FIG. 5 f, the dominant negative phenotype of the C22U/U40C CRE allele was eliminated when normal translation of the genome was blocked, arguing that it is not the CRE RNA alone, but some complex formed upon translation of viral proteins that inhibits the growth of other viruses in the same cell.

FIGS. 5A-F. RNA replication or translation requirements for dominance of mutant poliovirus alleles. (a) Schematic of mutant alleles mapped to the poliovirus genome assayed for dominance requirements. ΔGUA₃ is a genome lacking nucleotides G₇₄₁₈UAAA₇₄₂₂; RNAs that contain this 3′-non-coding region deletion are deficient for negative strand synthesis³⁶. The VP4-M1stop mutation changes the initial methionine of VP4 to a UAG stop codon. (b) Test of replication requirement for dominance of VP2-S243P genome. Co-transfections were performed in triplicate and are graphed as the average of replicate co-transfections with error bars to indicate standard error. (c) RNA replication requirements for 3D S291P. A mutant genome containing both the 3D-S291P and ΔGUA3 deletion was co-transfected with wild-type RNA and graphed as in b. The co-transfection of wild-type and a genome containing only the ΔGUA3 deletion is shown as a control. (d) RNA replication requirements for a mutant allele of the RNA replication protein primer, 3B-Y3H. Co-transfections were performed and graphed as in b, above. “ΔGUA₃ alone” indicates that no wild-type RNA was co-transfected, while “ΔGUA₃” indicates a co-transfection of that RNA with wild-type virus. “3B Y3H/ΔGUA₃” indicates the co-transfection of a non-replicating RNA that contains both mutations with wild-type RNA. No virus was detected at the highest dilution of the ΔGUA₃ virus alone, so the limit of detection is graphed. (e) RNA replication requirements for mutant CRE allele dominance. Co-transfections were performed as in b. “C22U/U40C/ΔGUA3” refers to the doubly mutant, non-replicating genome that contains the mutant CRE allele and 3′-non-coding region deletion. (f) Translation requirements for dominance of mutant CRE allele C22U/U40C. Co-transfections of the indicated genomes containing mutant alleles are shown as in b. “VP4-M1stop/C22U/U40C” is a non-translating genome that contains all indicated mutations.

Effects in Other Nonstructural Protein Coding Regions

The effect of mutations in the 2B, 2C, 3A, and 3C coding regions on co-transfected wild-type viral genomes is shown in Table 2. TABLE 2 Dominant inhibitor screen results of mutant 2B, 2C, 3A, and 3C alleles. PFU/ml Percentage (×10⁴) of control tRNA 23  100 ± 29% control* R2-Pvull 0.49 2 ± 1 2B F13S 2.5 11 ± 2  2B F17S 2.7 11 ± 5  2C L93P 18 76 ± 15 2C F242S 5.0 21 ± 13 2C F328S 10 44 ± 13 3A L8S 2.0 9 ± 2 3A F83S 10 44 ± 7  tRNA 24  100 ± 19% control* R2-Pvull 2.0 8 ± 2 3C-L70P 35 144 ± 32  3C-L102S 54 226 ± 31  3C-C147R 57 239 ± 26  2A-F133S 54 224 ± 35  *Independent sets of co-transfections are shown.

Of the three membrane-associated proteins 2B, 2C, and 3A, mutations in 2B were found to be more consistently dominant, although their suppression of wild-type growth was not as pronounced as that of the R2-PvuII control. Partially dominant mutations in the 2B coding region have been reported previously²³, although the mechanism of dominance remains unknown. Unlike mutations in 2A proteinase (FIG. 4), mutations in 3C proteinase were either recessive or gave rise to viruses that provided a helper function (Table 2).

Superinfections of Temperature-Sensitive and Wild-Type Polioviruses.

To test whether the locus- and allele-specific dominance of co-transfected genomes would also be observed with viable viruses, the ability of temperature-sensitive (ts) viruses (FIG. 6 a) to inhibit the growth of wild-type virus was monitored (Methods). First, cells were infected with ts mutant viruses in the presence of an RNA replication inhibitor, guanidine hydrochloride, to allow mutant viral proteins to accumulate to similar concentrations for each of the ts mutant viruses. At a later time point, guanidine was removed, wild-type virus was added, and a single cycle of viral growth continued at a semi-permissive temperature. The resulting virus stocks were then titered at a non-permissive temperature to quantify the yield of wild-type virus. As shown in FIG. 6 b, two mutant viruses with mutations in the 2A proteinase (S74P) and 2C NTPase (F28S) coding regions did not hinder wild-type growth. However, a virus that contained a mutation in the capsid coding region²⁴ (VP2-R76Q) reduced wild-type virus growth by more than 260-fold compared to wild-type growth alone (FIG. 6 c).

FIGS. 6A-C Superinfections of wild-type and temperature-sensitive polioviruses. HeLa cells were infected with the temperature-sensitive poliovirus indicated while blocking RNA synthesis as described in Materials and Methods. After mutant proteins were allowed to accumulate, wild-type poliovirus was added to HeLa cells as indicated, and the block to RNA synthesis released. Viral titers after a single cycle of infection are shown above for two separate experiments at the restrictive temperature. (a) Schematic of temperature-sensitive alleles used in superinfections. (b) Yield of wild-type virus following superinfection of cells that had accumulated protein from temperature-sensitive mutant viruses 2A-S74P or 2C-F28S (Table 3) as temperature sensitive viruses. (c) Yield of wild-type virus following superinfection of cell that had accumulated proteins from temperature-sensitive mutant viruses VP2-R76Q²⁴, 3D-F34L (Table 3), or 3D-T367I²⁵.

Two viruses with different mutations in the coding region for the viral RNA-dependent RNA polymerase, 3D-F34L and 3D-T367I (Table 3)²⁵, were tested for their ability to hinder wild-type growth. While 3D-T367I virus was not inhibitory, 3D-F34L exhibited a four-fold inhibition of wild-type virus growth, thus showing that dominance of mutant genomes differed between coding regions and between alleles of the same coding region. TABLE 3 Phenotypes of temperature-sensitive polioviruses generated by hydrophobic mutations. Phenotype PFU/μg PFU 39° C./ RNA 32.5° C. 39° C. 32.5° C. wild- 290 large plaque large plaque 1.4 type 2A- 190 very small none detected <0.07 S74P plaque 2C- 140 small plaque small plaque 0.05 F28S 3D- 990 small plaque very small 0.4 F34L plaque Summary of Dominant Negative Alleles

A genomic screen with poliovirus, a positive-strand RNA virus, was conducted to identify viral proteins that, when made nonfunctional by mutation, would dominantly interfere with the growth of co-transfected wild-type viral RNA genomes. These proteins should constitute ideal drug targets if the defects of the dominant alleles can be phenocopied by the antiviral compounds. Twenty-seven different genomes, each of which contained a single lethal mutation, were tested. Four classes of strongly dominant mutations were observed (Table 4). First, capsid mutations were dominant, presumably because nonfunctional mutant capsids co-assemble with wild-type capsids and render them nonfunctional. A second class of dominant genomes contained mutations in the poliovirus polymerase coding region; however, only two out of seven mutations in the polymerase coding region were strongly dominant. Poliovirus polymerase is known to oligomerize¹⁰; therefore, the mechanism of dominance is likely to be similar to that of the capsid alleles. A third class of strongly dominant mutations in poliovirus was found in an RNA structure, termed the CRE, that is required for generation of the protein primer for polioviral RNA synthesis. Finally, mutants that rendered the 2A proteinase of poliovirus inactive (L98P and C109R, FIG. 4) were dominant and profoundly inhibitory. The cleavage between VP1 and 2A coding regions within the viral polyprotein is made by 2A proteinase and reported to be intramolecular²¹, and would therefore be refractive to scission in a mutant proteinase even in the presence of mature, wild-type 2A proteinase encoded by coinfecting genomes. Uncleaved VP1-2A protein encoded by the mutant genomes may inhibit co-infecting genomes in the same way that mutant capsid protein does, by co-assembling with wild-type capsids and poisoning their function. To test this hypothesis, the effects of directly mutating the VP1-2A cleavage site were determined. These mutants were also dominant. TABLE 4 Summary of dominant negative alleles of poliovirus. Potential Mechanism Recessive or cis-Dominant Alleles^(a) 2A-F133S, VP4-M1stop Translation defect 2C NTPase (L93P) Potentially misfolded protein 3A (L24S) Potentially misfolded protein 3C proteinase (L70P, L102S, C147R) Potentially misfolded protein 3D polymerase (L107P, F246S, F296S) Potentially misfolded protein CRE “l.o.f.” (C13U/A16C/G19A/C22U/ Unfolded RNA G25A/A26U/G27C/A31G) Co-dominant Alleles^(b) 2B (F13S, F17S) 3D polymerase (F30S, F191S) 3A (L8S, F83S) Dominant Alleles^(c) Capsid (VP2-S1P, VP2-S243P, Chimeric encapsidation of VP3-L211S, VP1-L118P) wild-type genomes 2A proteinase (L98P, C109R) Defect in intramolecular cleavage yields toxic precursor 3D polymerase (S291P) Chimeric oligomers CRE (G19A/U40C, C22U/U40C) or Arrested RNA replication or VPg (3B-Y3H) priming complex ^(a)Greater than 80% of wild-type control. ^(b)Less than or equal to 80% of wild-type control and greater than R2-PvuII control. ^(c)Less than or equal to R2-PvuII control.

Example 2 Drug-Sensitive Virus Inhibits Growth of Drug-Resistant Virus, where the Drug Target is a Dominant Target

Methods

The methods are described in Example 1.

Results

The Presence of a Drug-Sensitive Virus Inhibits a Drug-Resistant Virus

The trans-acting, highly oligomeric nature of capsid proteins and the observed dominance of mutant capsid alleles suggested that a drug-resistant virus may inhibit a drug-sensitive virus if a particular drug targets a capsid protein. Disoxaril (WIN-51711) binds to the “canyon” residues of poliovirus virions, and through stabilization of the virion structure, prevents the uncoating of the viral genome after viral cell entry^(26,27). A mutation known to confer WIN resistance, VP1-I192F, was introduced into a cDNA encoding Sabin-3, the poliovirus serotype known to be most susceptible to the WIN-51711 (FIGS. 7 a,b)²⁸.

To mimic the situation in which a drug-resistant virus would appear in a cell infected with wild-type, drug-sensitive virus, co-infections of wild-type and WIN-resistant polioviruses were performed at a high multiplicity of infection (MOI) for the wild-type virus and a much lower MOI for the drug-resistant virus. As can be seen (FIGS. 7 a,b), the output of WIN-resistant virus was greatly reduced when grown in the presence of drug-sensitive virus, to 7% of the yield from a single infection. The effect was similar when the single-cycle co-infections were performed in the absence or presence of the selective agent. The observed dominance of the drug-sensitive genomes may be due to chimeric capsid formation, which rendered WIN-resistant genomes susceptible to the drug, being partially encapsidated by WIN-sensitive capsid proteins. An alternative explanation, however, was that RNA replication of the WIN-resistant virus was reduced by an unknown mechanism in the co-infection. To test this possibility explicitly, total RNA from all the infections in FIG. 7 a was subjected to RT-PCR. Differential digestion with a restriction enzyme was employed to determine the proportion of WIN-resistant genomes in each infection (FIGS. 7 c,d). As shown in FIG. 7 e, similar amounts of WIN-resistant viral RNA were present in both single and co-infections. The result argues that the reduction in WIN-resistant virus during co-infection was not due to a decrease in RNA replication, but to the formation of chimeric capsids that rendered the drug-resistant genome drug-sensitive.

FIGS. 7A-E. Co-infections of drug-sensitive and drug-resistant viruses. Viral infections were performed using either the poliovirus type-3 isolate “Fox” strain designated “WIN-S” or a WIN-51711-resistant derivative of the Sabin-3 strain containing a point mutation, VP1-I192F, labeled “WIN-R”. Infections were performed singly or as co-infections, for a single round of virus growth at the indicated MOIs. After virus adsorption, virus growth continued in the absence (a) or presence (b) of 2 μg/ml WIN-51711. To measure “total virus”, viral titers were determined in the absence of drug; WIN-R virus was measured by adding drug to virus stock dilutions and agar overlays. Percentages refer to the relative amounts of WIN-R virus when WIN-R virus from a single infection is defined as 100 percent with standard error measurements indicated by error bars. (c) Schematic of RT-PCR strategy used to measure ratio of WIN-S to WIN-R intracellular RNA. Total intracellular RNA was harvested after viral infection of HeLa cells grown in the absence of WIN at the MOIs indicated, subjected to RT-PCR using primers common to both WIN-S and WIN-R RNAs, and digested with a restriction enzyme, TfiI, to quantify the relative abundances of each RNA species. The asterisk (*) denotes the radiolabeled forward primer; the differential mobility of the restriction digested, radiolabeled RT-PCR product is used to distinguish WIN-R and WIN-S species in d,e. (d) Standard curve of in vitro transcribed WIN-R and WIN-S RNA. The indicated RNAs were produced from linearized cDNA templates in vitro and added to each RT-PCR reaction. The percent of total RT-PCR product that migrated as either WIN-S (white bars) or WIN-R (shaded bars) is shown for each reaction. The relative intensity of each product was quantified using a phosphorimaging plate and ImageQuant software (e) Quantitation of viral intracellular RNA from infected cells. RT-PCR reactions were performed as described in c using intracellular RNA harvested from the infections indicated in a. Each lane is labeled with the infection used in the RT-PCR reaction. “Mix” refers to a mixing of intracellular RNAs harvested from separate single infections of WIN-R and WIN-S viruses, while “co-infect.” refers to the intracellular RNA harvested from a co-infection of WIN-R and WIN-S viruses. The relative mobilities of WIN-R and WIN-S digested products are indicated in the left panel. To determine the relative abundance of WIN-S and WIN-R RNA species, the indicated bands were quantified using a phosphorimaging plate and ImageQuant software. The percent of total signal for WIN-S (white bars) or WIN-R (grey bars) are indicated in the right panel graph.

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While the present invention has been described with reference to the specific embodiments thereof, it should be understood by those skilled in the art that various changes may be made and equivalents may be substituted without departing from the true spirit and scope of the invention. In addition, many modifications may be made to adapt a particular situation, material, composition of matter, process, process step or steps, to the objective, spirit and scope of the present invention. All such modifications are intended to be within the scope of the claims appended hereto. 

1. A method of identifying a candidate anti-viral agent, the method comprising: a) culturing a mammalian cell in vitro, wherein the mammalian cell comprises: i) a parent RNA virus, wherein growth of the parent virus is inhibited by a test agent; and ii) a variant of the parent RNA virus, wherein growth of the variant RNA virus is resistant to the test agent; and b) determining the effect, if any, of parent virus growth on growth of the variant virus during at least one replicative cycle, wherein, when parent virus growth interferes with variant virus growth during at least one replicative cycle, the test agent is considered a candidate anti-viral agent.
 2. The method of claim 1, wherein, in the presence of the candidate anti-viral agent, parent suppresses growth of any drug-resistant variant virus for at least one replicative cycle.
 3. The method of claim 1, wherein the variant RNA virus genome comprises one or more changes in nucleotide sequence relative to the nucleotide sequence of the parent RNA virus, wherein the one or more changes in nucleotide sequence result in a change in the RNA and/or an encoded protein that inhibits growth of parent virus in a mammalian cell containing both parent virus and variant virus when the cell is cultured in the absence of the test agent.
 4. The method of claim 3, wherein the encoded protein is an oligomeric protein.
 5. The method of claim 3, wherein the encoded protein is a trans-acting protein.
 6. The method of claim 1, wherein the parent and the variant RNA viruses are positive-strand RNA viruses.
 7. The method of claim 1, wherein the parent and the variant RNA viruses are negative-strand RNA viruses.
 8. The method of claim 1, further comprising the step of selecting for the variant of the parent RNA virus.
 9. The method of claim 1, wherein the parent virus is a poliovirus, a retrovirus, or a hepatitis C virus.
 10. The method of claim 4, wherein the oligomeric protein is a capsid protein, a membrane-associated protein, or a polymerase.
 11. The method of claim 5, wherein the trans-acting protein is a protease that cleaves a polyprotein, a protein primer, or an RNA helicase. 